Studies with the RNA polymerase I. Factors affecting the binding of nucleic acid polymers to the enzyme

BIOCHIMICA ET BIOPHYSICA ACTA

458 BBA 96097

STUDIES

WITH THE RNA POLYMERASE

I. FACTORS

AFFECTING

POLYMERS

TO THE ENZYME

DONALD

D. ANTHONY*,

THE BINDING

EILEEN

ZESZOTEK

OF NUCLEIC ACID

AND

DAVID

A. GOLDTHWAIT’”

The Department of Pharmacology and the Department of Biochemistry, Case Western Reserve Univevsity, Cleveland, Ohio 44106 (U.S.A.) (Received

September

z?th,

1968)

SUMMARY

The association of RNA polymerase (nucleoside triphosphate: RNA nucleotidyltransferase, EC 2.7.7.6) with DNA or RNA as well as some of the factors which affect the enzyme-polymer complex have been examined. I. The 13-S form of the enzyme was bound to transfer RNA (tRNA). Binding of slightly more than I tRNA per enzyme molecule was observed. 2. An order of displacement for nucleic acid polymers was established by experiments which demonstrated the displacement of one polymer from the enzyme by a second polymer. The order was: single-stranded sonicated DNA 2 double-stranded sonicated DNA > single-stranded DNA > tRNA > native DNA. 3. Ionic strength affected the association of enzyme and polymer. Association was greatest at low ionic strength; in centrifugation experiments, (NH&SO, at 0.066 M caused a 90 o/0dissociation of the enzyme-polymer complex. In kinetic experiments with varying DNA concentrations, (NH&SO, produced noncompetitive inhibition. 4. Divalent cations decreased association. In the absence of divalent cations and in the presence of EDTA, zo--25 “/”more RNA or DNA was bound to the enzyme than in the presence of divalent cations. 5. Nucleoside triphosphates at elevated concentrations produced dissociation of the RNA-enzyme complex. The order for the displacement effect was ATP > GTP > UTP > CTP. Nucleoside triphosphates also produced either dissociation or aggregation of the DNA-enzyme complex. The order for this effect was GTP > ATP > CTP > UTP. A divalent cation was required for this nucleotide activity with both RNA and DNA. 6. Nucleoside triphosphates at elevated concentrations also inactivated the enzyme. The order of activity for this effect was GTP > ATP > CTP > UTP. 7. The effects of the nucleotides lead to a prediction that a site exists on the RNA polymerase which interacts preferentially with the purine nucleotides. It is suggested that this may be the binding site for the 5’-terminal nucleoside triphosphate. American Cancer Society Faculty Research Associate. ** National Institutes of Health Research Career Investigator ??

Rioc’lim.

Biophjw. Acta,

174 (1969) 458-475

Fellowship.

STUDIES

WITH

THE

RNA

POLYMERASE

459

1

INTRODUCTION

The formation of a complex between DNA and the RNA polymerase (nucleosidetriphosphate: RNA nucleotidyltransferase, EC 2.7.7.6), which did not dissociate during the synthesis of RNA, was first demonstrated by BREMER ANDKONRAD~.They also showed that the RNA which was synthesized remained with this DNAxnzyme complex. A number of binding sites on the polymerase are required for this complicated reaction. In a previous paper2, preliminary kinetic evidence was presented for two different nucleoside triphosphate binding sites. Another site on the enzyme is involved in the binding of the template. This site is not absolutely specific for deoxyribonucleotide polymers as shown in a number of studies of the ability of the RN-4 polymerase to copy ribonucleotide polymers 3- B. Further evidence for RNA binding is the inhibition of DNA-directed RNA synthesis caused either by the addition of RNA polymers, particularly transfer RNA (tRNA), or by the RNA synthesized during the reaction’-13. This inhibition is most likely due to competition between the polymers for the template binding site. Thus a template binding site exists, to which either deoxyribonucleotide or ribonucleotide polymers are bound. The experiments presented in this paper were designed to obtain more specific information about factors which affect this binding site. Data concerning the affinity of different nucleic acid polymers for this binding site, and the effect of ionic strength, divalent cations, and nucleoside triphosphates on the stability of the polymer-enzyme complex are presented. A preliminary communication of this material has appeared14.

EXPERIMENTAL

PROCEDURE

Materials and methods. RNA-32P-labeled

tRNA was prepared from Escherichia by the procedure of STENT AND FUERS+~. The bacteria were grown from 108 to 10~ cells/ml in IOO ml of minimal mediumIs which contained 4 mC of 32Pi. After centrifugation at room temperature, the cells were resuspended in IOO ml of minimal nonradioactive medium and were allowed to grow for one additional generation. IOO ml of cells, grown in a similar fashion but without 32Pi, were then added, and all cells were washed once in 0.001 M Tris-HCI buffer (pH 7.1) containing 0.01 M magnesium acetate. tRNA was then extracted according to the procedure of ZURAY”. This RNA preparation was purified further by fractionation on a methylated albumin column according to the method of MANDELLand HERSHEY~~as modified by SUEOKA~~.The final preparation migrated in a sucrose density gradient with a coefficient of 4.0 S relative to 16- and 23-S E. coli ribosomal RNA. Unlabeled E. coli tRNA was prepared by the same procedure or was purchased from the General Biochemical Corp. and chromatographed on a methylated albumin column. DNA. Calf thymus DNA was prepared according to the procedure of HURST~O, and unlabeled and [14C]thymine-labeled T4 DNA by the method of MANDELLAND HERSHEY’~. The different DNA’s were denatured by heating to 100’ for IO min or by adjustment of the pH of the solution to 11.5 for IO set in a N, atmosphere21. DNA of bacteriophage T4 was sonicated at o” with a Branson Sonifier (Model LS-75). The coli SW-1485

Biochim.

Bioph.ys.

Acta,

174

(1969)

458-475

1). D. ASTHOKY

460 average sedimentation ments was 7.8.

coefficient

of the sonicated

preparations

et al.

used in these expel--

Formation and sedimentation of the polymer-ewyme complex. KKA or DNA was incubated with the enzyme at o” in a mixture containing 50 mM Tris-HCl (pH 7.5) and 5.4 mM mercaptoethanol, in a final volume of 0.2 ml. Divalent cations, EDTA and nucleotides were also added as indicated. The sequence of addition was buffer, mercaptoethanol, and divalent cations, followed by enzyme and then RNA or DNA. In experiments designed to demonstrate competition between nucleic acid polymers, the enzyme and the first polymer were incubated either 5 min or IO min at o” before addition of the second polymer. For the investigation of the effect of a nucleoside triphosphate on the RNA- or DNA-enzyme complex, the incubation mixture which contained polymer and enzyme was incubated for 5 min at o0 before addition of the nucleotide. (Formation of the nucleic acid-enzyme complex was shown in other experiments to be complete within 5 min.) After the final addition, mixtures were incubated for IO min at 0' and then layered on a 4.4-ml linear 5-20 o/osucrose gradient containing 50 mM Tris-HCl (pH 7.5) and divalent cations or EDTA as indicated. The components were separated by centrifugation at 25 ooo rev./min for 15-16 h in a SW39 rotor in a Spinco Model L ultracentrifuge at 5” unless otherwise noted; 32 or 33 fractions were collected from each tube. Assay of components. 14C-labeled T, DNA or 32P-labeled tRNA, either free or as a nucleic acid-enzyme complex, was assayed by counting 50 ~1 of each fraction from the sucrose gradient in a Packard Tri-Carb liquid scintillation counter. In some cases nucleic acid was assayed by measuring A26,, mp. RNA polymerase, free or bound, was assayed by addition of 20 ,ul of each fraction to the standard assay mixture described previously2. Units of enzyme activity recovered from sucrose gradients were approx. 60-65 y. of the total units added to each gradient. The recovery of radioactive RNA approached IOO o/O,and the i4C-labeled DNA recovery was 85-100 y0 except in experiments where aggregation occurred and a fraction of the DNA sedimented to the bottom of the tube. Enzyme. RNA polymerase was purified from E. coli by a variation* of the method of MAITRA AND HURWITZ 22. The purified enzyme had a specific activity of ~OOO6000 units/mg with calf thymus DNA as a template where a unit was defined as the amount of enzyme required for the incorporation of I nmole of labeled nucleoside monophosphate in 60 min at 37’. The enzyme was stored at 2’ in 30 y0 (NH,),SO,. Miscellaneous. Nucleotides and other materials are described in a previous paper2 or in the legends to figures and tables. The concentration of tRNA was determined (ref. 23). The concentration of DNA solutions was determined by by its A,,, mp A 26,,mp (ref. 24) and by a modification of the diphenylamine reactionz5. The degree of denaturation of DNA preparations was determined by measurement of absorbancemelting curves in a Gilford spectrophotometer. Denatured DNA in 0.015 M NaCl, 0.0015 M sodium citrate had a hyperchromic shift of less than 5 o/Oin the temperature range from 40’ to 96”.

RESULTS Biding *

of RNA

Polymerase to tRNA

J. HURWITZ, personal communication.

Riochim.

Riofihys.

Acta,

174 (1969) 458-475

RNA polymerase

forms a complex

with

STUDIES

WITH THE

RNA

POLYMERASE.

1

461

either RNA or DNA. Since tRNA represents a family of molecules with a fairly uniform molecular weight (approx. 25 ooo), initial studies of binding between polymer and enzyme were done with this RNA. Fig. IA shows data obtained from a sucrose density gradient centrifugation experiment in which an excess of E. coli 32P-labeled tRNA was mixed with the RNA polymerase purified from E. coli. The free tRNA peak was in Tube 26, while the peak of the tRNA-enzyme complex was in Tube 15. The activity of the RNA polymerase coincided almost exactly with the radioactivity of the RNA in the complex. The sedimentation coefficient of free RNA polymerase was 13.4 relative to alkaline phosphatase in a 5-20 oh sucrose gradient which contained 0.05 MTris-HCl (pH 7.5) plzts 0.001 M EDTA. The sedimentation coefficient of the RNA-enzyme complex was 14.4 relative to alkaline phosphatase. Storage of the enzyme in 30 yb (NH,),SO, was probably responsible for the low S value obtained in a gradient of low ionic strength. In some experiments a minor component which amounted to 10-20 ‘$b of the total complex was observed which had a sedimentation coefficient of approx. 21. The presence of this fraction was not constant, and the conditions required for its observation have not been studied. Thus it is clear that the 13-S form of the enzyme, which corresponds to a molecular weight of 370 ooo (ref. 22) or 440 ooo (ref. 26), binds to tRNA. Binding of this form of the enzyme has been noted by others11v27. The stoichiometry of binding of tRNA to the RNA polymerase was determined, and the results are indicated in Table I. The peaks of the RNA-enzyme complex had relative S values of approx. 14. In Expt. I, excess RNA relative to the enzyme was added. Free RNA as well as the RNA-enzyme complex was observed. In Expt. 2, addition of 2 times the amount of RNA did not increase the amount of the complex significantly, while in Expt. 3 a doubling of the enzyme concentration plus a 3-fold increase in RNA relative to that in Expt. I resulted in a doubling of the complex observed. Based on the assumption of a molecular weight of 25 ooo for the tRNA and 370 ooo for the enzyme, the molar ratio of RNA to enzyme in the complex was 1.2-1.4 assuming the enzyme was pure. The specific activity of the enzyme preparation used in this experiment was 4900 nmoles of the radioactive nucleotide incorporated per h per mg of protein at 37O with calf thymus DNA as the template. This was 80 y0 of the specific activity of our most pure enzyme preparation and 66 oh of that obtained by RICHARDSoNB6with T7 DNA as a template. It is possible that two tRNA molecules were bound to the enzyme. It is difficult to rule out the presence of enzymatitally inactive RNA polymerase protein which retained its capacity to bind to polymer. One reason that the ratio of RNA to enzyme in the complex was higher in these experiments than the ratio observed by others may be the absence of divalent cations, an effect which will be discussed later. (In other binding experiments reported in this paper, the ratio of RNA to enzyme required for saturation was lower. In these cases either the specific activity of the enzyme was less, often due to gradual inactivation on storage, or the ionic conditions of both the reactions and the sucrose gradients were different.) Binding of RNA polymerase to DNA. The binding of the enzyme to sonicated 14C-labeled T4 DNA is shown in Fig. IB. The peak of excess free DNA was in Tube 22 while the DNA-enzyme complex was present in a broad peak which was spread through the lower part of the gradient. Enzyme activity coincided moderately well Riochlm.

Biophys.

Acta,

174 (1969)

458-475

462

I).

4

6

12

I).

ANTHONY ?t al.

1620242632

FRACTION

NUMBER

FRACTION

NUMBER

Fig. I. Sedimentation of RNA- and DNA-enzyme complexes in sucrose density gradients. A. tRNA-enzyme. Formation of the complex: 224pg of E. coli RNA polymerase were added to a mixture containing 50 mM Tris-HCl (pH 7.5), 2 mM MnCl,, and 8 mM Cl:. After 5 min at o’, 7.74 pg of 32P-labeled tRNA were added. This mixture, in a final volume of 0.2 ml, was then incubated for an additional IO min at 0”. Centrifugation and assay: the reaction mixture was layered on 4.4 ml of a 5-20 y/o sucrose density gradient containing 50 mM Tris-HCl (pH7.5), 2 mM MnCl,, and 8 mM MgCl,. The components were centrifuged, and fractions were collected and assayed for sedimentation of the labeled tRNA as described in Materials and methods. For enzyme activity zo,nl of each fraction were preincubated in a standard reaction mixture containing 50 mM potassium maleate (pH 7.5), 2 mM MnCl,, 8 mM MgCl,, 5 mM mercaptoethanol, and 15 ,ug of single-stranded T4 DNA for ro min at 0’. The reactions were then started by the addition of 0.4 mM ATP, GTP, and UTP and 0.2 mM 3H-labeled CTP (1694 counts/mm per nmole). Incubations were for 20 min at 38”. The RNA in o.3-ml aliquots was precipitated, washed, and counted as described2. The maximum radioactivity of tRNA with an S value of 4 was in Fraction 27. B. DNA-enzyme. The complex was formed as in A except that 168 ,ng of enzyme were incubated with 33 pig of heat-denatured, sonicated ‘K-labeled T4 DNA. The solution was then layered on a sucrose gradient and centrifuged as in A. Sedimentation of the labeled DNA was similar to A, and enzyme activity was assayed as in A except that 25 pg of calf thymus DNA were added, there was no preincubation, and the substrate was 32P-labeled UTP (1500 counts/ min per nmole) Fig. 2. Relative displacement of tRNA from the tRNA-enzyme complex by different preparations of DNA. A. The polymer-enzyme complex was formed in a. final volume of 0.2 ml containing 50 mM Tris-HCl (pH 7.5), 2 mM MnCl,, and 8 mM MgCl,. Expt. I contained only 22 pg of sonicated r%labeled T4 DNA. In Expt. 2, I 12 pg of RNA polymerase and 2.6 ,ng of 32P-labeled tRNA were incubated IO min at o” to form tRNA-enzyme complex. In Expt. 3, the tRNA-enzyme complex was formed as in 2 except that after 10 min at on, 22 pg of sonicated W-labeled Tq DNA were added, and the solution was incubated an additional IO min at 0’. The solutions were then layered on 4.4 ml of a linear 5-20 ye sucrose density gradient containing buffer and divalent cations as in the incubation. Centrifugation and assay for labeled polymer was as described in Materials and methods. B. Experimental conditions were as described in A, Expt. 3, except T4 DNA, instead of the sonicated preparation, were that 23 pg of single-stranded i%labeled added to the “ZP-labeled tRNA_enzyme complex. C. Experimental conditions were again as in A, Expt. 3, axcept that 23 ,ug of native W-labeled T4 DNA, instead of the sonicated preparation, were added to the 3ZP-labeled tKNA-enzyme complex.

with the DNA radioactivity that sedimented beyond the free DNA. The lack of exact correlation between enzyme activity and the W-labeled DNA in the complex may Biochim.

Biophys.

Acfa,

174 (Ig6g)

458-475

STUDIES

TABLE

WITH THE

RNA

POLYMERASE.

1

463

I

s’rorcnIoMErRY

OF BINDING BETWEEN

RNA

POLYMERASE

AND

E.

coii

tRNA

tRNA and enzyme, in the concentrations indicated in the table, were incubated in 0.05 M TrisHCl (pH 7.5), 5.4 mM mercaptoethanol at o0 for 5 min, to form tRNA-enzyme complex, and then centrifuged in a sucrose gradient. Details of the gradient, measurement of the position of labeled tRNA (complex and free) and the enzyme assay are described in Materials and methods. Gradients contained 0.001 M EDTA, and no divalent cation was present either during formation of the complex or in the sucrose gradient. The percent of tRNA, both free and in the complex, was determined by mea.suring the area under the two peaks obtained by plotting counts 11s.fractions. The molecular weight of E. coli tRNA was assumed to be 25 ooo, and that of the RNA polymerase 370 ooc (ref. 22).

Expt. No.

EVZZYWU? added (f%)

tRhTA added (Icg)

tRNA in enzyme complex

Free tRNA (Pl)

tRNA/ enzyme wt. ratio

tRNA / enzyme molar ratio

0.06 0.06 0.07

I.3 I.4

(H) 28 28 56

I 2

3

3.8 7.6 II.4

1.6 I.7 3.9

2.2

5.8 7.4

I.2

be due to the presence of multiple binding sites on each DNA molecule; in the peak is very likely a consequence of this.

the broadness

order of binding of tRNA and DNA to the RNA polymerase. A series of was done which demonstrated that the binding of the RNA polymerase to different polynucleotides was reversible, and that an order of displacement of different polymers from the enzyme existed. The first experiment, illustrated in Fig. 2A, showed that sonicated doublestranded DNA would displace tRNA from the enzyme. Curve I shows the position of free T4 sonicated 14C-labeled DNA in a sucrose density gradient centrifugation experiment. The peak had a sedimentation coefficient of 7.8 relative to tRNA. Curve 2 shows the tRNA-enzyme complex and free tRNA obtained from a second tube. Most of the tRNA was present as an RNA-enzyme complex with its peak in Fraction 12; a small amount of excess free RNA with a peak in Fraction 25 was also present. For the data illustrated in Fig. 2, Curves 3a and b, 32P-labeled tRNA and enzyme were incubated for IO min at 0”, then sonicated r4C-labeled DNA was added and the incubation was continued at 0’ for an additional IO min prior to addition of the mixture to the centrifuge tube. Curve 3a represents the “C-labeled DNA with a variable amount of bound enzyme per DNA molecule. The low 14C peak in Fraction 22 represents small fragments of free DNA. Curve 3b shows the 32P-labeled tRNA pattern in the same experiment. This RNA was no longer bound to the enzyme. Thus, sonicated DNA displaced tRNA from the RNA polymerase. Single-stranded DNA, not sheared by sonication, was also able to displace tRNA from the enzyme. Incubation of 32P-labeled tRNA and enzyme followed by addition of single-stranded i4C-labeled DNA and further incubation prior to centrifugation, produced the patterns shown in Fig. 2B. The RNA was almost totally displaced from the RNA-enzyme complex while most of the high molecular weight single-stranded r4C-labeled DNA sedimented presumably with the enzyme to the bottom of the tube. A control without added DNA showed 80 yO of the RNA present as a complex with the enzyme. When sonicated double-stranded DNA was converted Relative

experiments

Biochim.

Riophys.

Acta.

174 (rg6g)

458-475

D. D. ANTHONY Pt al.

464

to single-stranded DNA, this also displaced the RNA completely from the RNA-enzyme complex. Double-stranded i4C-labeled T4 DNA, not sheared by sonication, was unable results shown in to displace 32P-labeled tRNA from the enzyme. The experimental Fig. ZC were obtained by additions in an order similar to those in Fig. zB. The percent of total 32P-labeled tRNA observed in the complex with the peak in Fraction 18 was 79 7; of that observed in a control tube without addition of DNA. In Fig. 2C, the high molecular weight DNA sedimented to the bottom of the tube. Thus, doublestranded high molecular weight DNA did not displace significantly the tRNA from the enzyme under these conditions. In a similar experiment, when this X-labeled DNA was first incubated with the enzyme and then 32P-labeled tRNA was added, the DNA was displaced by the tRNA. To provide direct evidence that sonicated DNA actually reversed the association between RNA and enzyme, the 32P-labeled tRNA_enzyme complex was first isolated from a sucrose gradient centrifugation experiment; this complex was then incubated with either sonicated double-stranded or untreated double-stranded T4 DNA, and the mixtures were recentrifuged. The results indicated that the tRNA in the complex was displaced by sonicated DNA, but not by the high molecular weight DNA. It is apparent that sonicated DNA with an average sedimentation coefficient of approx. 7.8 had properties similar to single-stranded DNA. An order of displacement of these nucleic acid polymers for an enzyme polymer was thus established as sonicated DNA 2 single-stranded DNA > tRNA > double-stranded DNA. Further evidence that tRNA was able to dissociate the native T4 DNA-enzyme complex was obtained in experiments (Table II, Tubes 1-3) which employed the membrane filter technique originally developed by JONES AND BERGEN.These data show that almost two thirds of the complex formed between T4 DNA and enzyme was dissociated in the presence of 0.066 M (NH,),SO, when tRNA was added. This TABLE

II

THEEFFECT PLEX

OF tRNA,

NUCLEOTIDES

AND

(NH&SO,

ON THE **C-LABELED

DNA-ENZYME

COM-

'K-Labeled T4 DNA-enzyme complex was formed in a final volume of 0.5 mlcontaining 50 mM potassium maleate (pH 7.5), 2 mM MnCl,, 8 mM MgCl,, 5.4 mM mercaptoethanol, 15 ,ug of RNA polymerase, II pg of W-labeled Tq DNA and 0.4 mM nucleoside triphosphates as indicated. Following 2 min at 28’. a final concentration of 0.066 M (NH,)SO, was added where indicated. After an additional 2.5 min, 8g pg of tRNA were added as indicated. Reactions were incubated 15 min at z8”, and o.4-ml aliquots were filtered on Millipore membranes, washed and counted.

Tube No.

o/0of control DNA retained on

Order of additions

filter

Expt. Enzyme, DNA, (NH,),SO, Enzyme, DNA, (NH&SO,, tRNA Enzyme, DNA, tRNA Enzyme ,DNA Enzyme, DNA, ATP, GTP, UTP, CTP Enzyme, DNA, ATP, GTP, (NH&SO,, Enzyme, DNA, UTP, CTP, (NH,),SO,. Biochim.

Bioehys.

A&,

174 (v&g)

458-475

100

tRNA tRNA

4’ 55 ‘04 89 59 46

I

lspt.

100

35 76 66 58 31

2

STUDIES WITH THE

RNA

POLYMERASE 1

4%

dissociation occurred less readily in the absence of 0.066 M (NH,),SO,. This increased dissociation in the presence of (NH,),SO, may be due to the effect of ionic strength on the dissociation of the DNA-enzyme complex. This effect will be discussed further. The T4 DNAenzyme complex was dissociated less readily by tRNA if RNA synthesis was initiated either with the four nucleoside triphosphates or with ATP plus GTP (Table II, Tubes 5 and 6); UTP plus CTP had no effect. It is probable that the varying degree of reversibility noted by others in binding experiments10,r2 and kinetic experiments7sg is a function of ionic strength. The dissociation of the DNAenzyme complex by tRNA and the stability of the DNA-enzyme complex once RNA synthesis was initiated, is in accord with the nature of the tRNA inhibition observed by

others7,%1%12,29.

Effect of dieerent DNA preparations ed by tRNA

and on the tRNA-enzyme

on the inhibition of RNA polymerase

caus-

complex. Results

of the previous experiments indicated that the order of displacement of nucleic acid polymers for the polymerenzyme complex was: sonicated DNA z single-stranded DNA > tRNA > doublestranded DNA. These results suggested that tRNA and DNA compete for a polymer binding site on the enzyme. Experiments were then designed to study the ability of different DNA preparations to dissociate the RNA-enzyme complex. This dissociation was measured either by the degree to which these DNA preparations could reverse the inhibition of RNA synthesis caused by tRNA, or by the degree to which these DNA preparations could cause dissociation of the RNA-enzyme complex in centrifugation experiments. The results of these two experimental approaches are shown in Table III. Although the conditions are somewhat different in the two types of experiments, the order of displacement of different DNA preparations relative to RNA is the same. In both types of experiments shown in Table III, the tRNA_enzyme comTABLE

III

EFFECT OF ON

THE

DIFFERENT

tRN?-ENZYME

DNA’s

ON

THE

tRNA

INHIBITION

OF

THE

E. di

RNA

POLYMERASE

AND

COMPLEX

analysis. Reaction mixtures, 0.5 ml in volume, which contained 50 mM potassium maleate (pH 7.5), 5.4 mM mercaptoethanol, 0.4 mM ATP, GTP, and CTP; 0.1 mM 3aP-labeled UTP (1947 counts/min per nmole) and 14 pg of enzyme were preincubated IO min at o0 with or without 80 pg of E. coli tRNA. The reactions were then started by the addition of the DNA preparation indicated in the table; native T4 DNA, 5.6 pg; denatured T4 DNA, 6.2 ,ug; sonicated T4 DNA, 5.8 pg or sonicated denatured T4 DNA, 4.9 pg. Incubations were for IO min at 38”. Reactions were terminated and precipitates were washed and counted as described*. Centrifugation analysis. Experimental conditions were as described in Fig. zA, Expt. 3. tRNA-enzyme complex was formed by the addition of 2.6 pg saP labeled tRNA to IIZ pg of enzyme. After IO min at o0 the indicated preparation of DNA was added; native T4 DNA, II ,ug; denatured DNA, 12 ,ug; sonicated DNA I I ,ug or sonicated denatured DNA, 9 pg. After an additional 10 min at o’, the solutions were centrifuged and assayed as described in Materials and methods. The per cent of free and bound tRNA was determined by planimetry. _ T4 DN24 Kinetics analysis. Centrifzkjiation analysis. preparation Inhibition by RNA-enzyme complex renzainin~ affer tRNA (%) DNA addition (%) _ Native 90 79 Denatured 21 70 Sonicated 67 4 Sonicated and denatured 0 53 Kinetics

Biorhim.

Biophys.

Acta,

174 (1969) 458-475

466

D. I).ANTHONY

plex was formed before addition of the DNA preparation.

et al.

The amount of each DNA

preparation added in the kinetic experiments was similar. This condition also applied to the centrifugation experiments (see legend to Table III). The concentration of tRNA used in the kinetic experiments was well in excess of the concentration required to produce the maximum degree of inhibition (90 o/0) which could be obtained when native DNA was used as a template. From these data (Table III), mers from the RNA polymerase

the following order of displacement

was obtained:

denatured

of DNA poly-

sonicated DNA > sonicated

DNA > denatured DNA > native DNA. This data suggests that the enzyme has a higher affinity for the ends of DNA fragments than for areas in the middle of a single strand.

BERG et a1.30 have reported

The affinity

direct evidence for this type of terminal

of the enzyme for tRNA

The displacement

again appeared

greater

binding.

than for native DNA.

of one nucleic acid from an enzyme-polymer

complex

by a

second nucleic acid suggested that there was some form of competition for an enzyme site or sites. Further evidence for this competition was obtained. The synthesis of RNA by the RNA polymerase was examined as a function of varying DNA concentration with and without different fixed concentrations of tRNA. When the reaction was started by the addition of enzyme, and the results were plotted in the double reciprocal form, a competitive type of inhibition was observed. This effect has been observed by others7sg,10,12,2g. This evidence and the polymer displacement

data discussed

above both suggest that DNA and RNA compete for a similar site or sites on the enzyme. It should be emphasized that because of the size of these polymers, multiple binding sites cannot be ruled out. It is clear from the experiments

of others7,g,10,12,2g

and the results reported in this paper that the order of addition of DNA and RNA, the form of DNA, the ionic strength, and other factors all contribute to the degree of competitive been

or noncompetitive

inhibition

which can be observed.

The effect of (NH,),SO, 012 the dissociation of the RNA-enzyme complex. It has reported12,14,31 that the DNA-enzyme complex can be dissociated by high

ionic strength. The relative

Similar

results have been observed

dissociation

of the RNA-enzyme

for the RNA-enzyme

complex

0.044, or 0.066 M (NH,),SO, when present throughout dient is shown in Fig. 3. These molarities caused dissociation

0.022,

complex14.

caused by the presence

of

a sucrose density graof 52, 66, and 90 96 of

the complex, respectively. Complete dissociation was observed at 0.133 M (NH,),SO,. This dissociation had no requirement for a divalent cation. A sharp band of active enzyme,

separate

from free tRNA,

grated in the gradient

containing

was always recovered in the gradient. This miwith a sedimentation coefficient of 11.7

(NH,),SO,

relative to alkaline phosphatase. The reason experiments in which NH&l was employed in the dissociation of the RNA-enzyme complex was observed with 0.28 M NH&l. Thus, relativity low levels of (NH,),SO* in a centrifugal field (Fig. 3). However, 0.066

for the lower S value is not clear. In place of (NH,),SO,, similar effects on were obtained. Complete dissociation produced dissociation of the complex M (NHI),SO,, which produced 90 s/,

dissociation in these centrifugation experiments, did not significantly decrease the amount of native T4 DNA-enzyme complex when measured by the membrane technique. In the experiments shown in Table II, 0.066 M (NH,),SOI was present for 17.5 min after the DNA-enzyme complex had been formed, yet the amount of complex observed was the same as in the absence of (NH,),SO, (Tube I vs. Tube 4). Thus, Biochim.

Biophys.

Acta,

174

(1969)

458-475

STUDIES WITH ‘IHE

RNA POLYMERASE. 1

467

4 FRACTION

NUMBER

8 12 16 2O 24 28 FRACTION NUMBER

32

Fig. 3. Dissociation of the tRNA-enzyme complex due to (NH,),SO,. The tRNA-enzyme complex was formed in a final volume of 0.2 ml which contained 50 mM Tris-HCl (pH 7.5), 0.001 M EDTh, I 12 ,ug of enzyme, and 4.8 pg of 32P-labeled tRNA. After IO min at 0”. the solution was added to 4.4 ml of a linear 5-20 o/0 sucrose gradient which contained buffer and EDTA as in the incubation and the (NH,,),SO,i concentration indicated in the figure. Centrifugation and assay of the labeled tRNA, bound and free, was as described in Materials and methods. The maximum radioactivity of tRNA with an S value of 4 was observed in Fraction 26. Fig. 4. Dissociation of the tRNA-enzyme complex due to ATP. The tRNA-enzyme complex was formed in a final volume of 0.2 ml which contained 50 mM Tris-HCl (pH 7.5). 2.0 mM MnCl,, 8 mM MgCl,, 42 ,ug of RNA polymerase, and 0.95 ,~g 32P-labeled tRNA. After 5 min at o’, the concentration of ATP indicated was added. The solution was incubated an additional 10 min at o0 and then layered onto a 5-20 O/e sucrose density gradient which contained buffer and divalent cations as in the incubation. Centrifugation and assay of the labeled tRNA, bound and free, and methods. The maximum radioactivity of tRNA with an were as described in Materials S value of 4 was observed in Fraction 26 or 27.

with 0.066 M (NH&SO,, association was noted with DNA in the absence of a centrifugal field. However, the data in Table II (Expts. z and 3) also suggest that 0.066 M (NH,),SO, does increase the dissociation of the DNA-enzyme complex in the absence of a centrifugal field if tRNA is present. These data suggest that minimal dissociation of the DNA-enzyme complex can occur with 0.066 M (NH,),SO,, but that it is observed only if the equilibrium of the system is disturbed, for example by another binding agent such as tRNA or by a centrifugal field acting over a period of hours. Another factor which could be partially responsible for these results is the requirement for the binding of multiple enzyme molecules to DNA in order to observe retention of DNA on the filter32. Since (NH&SO, produced dissociation of the enzyme-DNA complex, it was possible that it acted directly on the polymer binding site. Kinetic experiments were therefore done to determine whether (NH&SO, competed with DNA to inhibit RNA synthesis. RNA synthesis was examined as a function of different DNA concentrations in the absence and in the presence of 0.05 and 0.1 M (NH&SO,. Reciprocal plots of the data indicated that this inhibition was noncompetitive. At 0.2 M (NH,),SO, the kinetics did approach more nearly a competitive pattern of inhibition. Other preliminary experiments indicated that in addition to affecting the dissociation of the Riochim.

Biophys.

Acta,

174 (1969)

458-475

468

11.1).ANTHONY

et cd.

DNA-enzyme complex, (NH,)&O, also inhibited the process of polymerization. This could be a reason for the mixed type of inhibition. The effect of divalent metals on the association of the enzyme with RXA and DLVA.

A divalent cation is not required for binding between a nucleic acid polymer and the enzyme (Table IV). When either DNA or RNA was incubated with the enzyme in

TABLE

IV

THE

EFFECT

AND

THE

OF DIVALENT

ABILITY

OF

ATP

CATIONS

ON THE

TO DISSOCIATE

AMOUNT THESE

OF

DNA

OR RN&ENZYME

COMPLEX

FORMED

COMPLEXES

The RNA-enzyme complex was formed in a final volume of 0.2 ml containing 50 mM Tris-HCl (pH 7.5); reaction mixtures without divalent cation contained 0.001 M EDTA; reaction mixtures with divalent cation contained 2 mM MnCI, and 8 mM MgCl,. RNA polymerase (42 ,~g) was added followed by 1.91 pg aaP-labeled tRNA. After 5 min at oO, 2.5 mM ATP was added to those experiments designated +ATP. Following an additional IO min at o’, solutions were layered on 4.4 ml of a linear 5-20 oh sucrose gradient containing buffer and divalent cation or EDTA as in the incubation. The gradients were centrifuged and then analyzed as in Materials and methods. The DNA-enzyme complex was formed, centrifuged, and assayed as the RNA-enzyme complexes except that 63 yg of enzyme was incubated with 7.3 pg of sonicated r%labeled DNA for 5 min The amount of at 0’. and then ATP (6 mM) was added to those solutions designated +ATP. polymer, bound of free, was determined by planimetry. In the absence of divalent cation and ATP, 82 yO of the tRNA and 79 yO of the T4 DNA were bound to protein. The data in the table are relative to these values. Relative anmunt of nucleic acid in nucleic acid-enzvme complex

Covnplex

- divalent cat&s -._____.. ~~ -ATP +ATP RNA-enzyme DNA-enzyme

+ divalent cations -.4TP

+ATP 0.07 0.04

I.0

0.98

0.80

1.0

I.0

0.76

the presence or absence of divalent cation and the amount of the complex was examined by sucrose density gradient centrifugation, the presence of the cation caused a decrease in the amount of polymer which was bound by the enzyme. 20 “/o less tRNA and 24 y0 less DNA were bound to the same amount of enzyme when Mg2+ and Mn2+ were present than when they were replaced by IO-~ M EDTA. Either magnesium or manganese alone decreased the amount of nucleic acid bound. This effect of divalent metals on polymer binding has also been observed by STERNBERGER AND

STEVE.NS~~.

The effect of ndeoside triphosphates on the tRNA-enzyme complex. The complex formed between RNA and the enzyme can be dissociated by high concentrations of nucleoside triphosphates. The effect of ATP is shown in Table IV. When 2.5 mM ATP was added to the RNA-enzyme complex in the presence of divalent cations and the mixture was examined by centrifugation in a sucrose density gradient, dissociation of the complex occurred. The data in Table IV indicate that divalent cation is required for this dissociation. In additional experiments it was found that either divalent cation alone had a similar effect. This is in contrast to the dissociation by (NH,),SO, which does not require a divalent cation. Typical centrifugation patterns which illustrate the dissociation effect of ATP Biochim.

Biophys.

Acta,

174 (1969) 458-475

STUDIES WITH THE

RNA POLYMERASE. 1

469

are shown in Fig. 4. In the control experiment without nucleoside triphosphates, almost all of the tRNA was present as a complex with the enzyme. The decrease in RNA in the complex when 0.75 and 1.25 mM ATP was present in the preincubation was 54 and 66 O$,,respectively. At 2.5 mM ATP (not shown) only 15 y0 of the tRNA remained bound to the enzyme. The concentration of nucleoside triphosphates required to cause dissociation of the RNA-enzyme complex varied with the enzyme preparation, but a specific order of nucleoside triphosphates was always observed (Table V). ATP was most TABLE

V

THE EFFECT OF ASIXAND tRNA

NUCLEOSIDE

TRIPHosPHATEs

ON THE

COMPLEX

FORMED

BETWEEN

RNA POLYMER-

62 pg of enzyme and 2.58 pug 32P-labeled tRNA were incubated in a final volume of 0.2 ml under conditions described in the legend to Fig. IA. After 5 min at o’, a single nucleoside triphosphate was added as indicated, and the mixture was incubated an additional ro min at 0”. It was then layered on a 4.4-ml linear 5--20 y0 sucrose gradient, centrifuged, and assayed as in Materials and methods. The per cent tRNA, bound and free, was determined by planimetry. Nucleotide addition

tRNA in the complex (%) ~_ 1.2 mM nucleotide

None .4TP GTP HTTP CTP

86 r9

0

:: 84

30 39 75

2.4 mM nucleotide

effective in causing dissociation of the RNA enzyme complex followed by GTP and then UTP. At the concentrations studied, CTP had little effect. Other compounds were tested for their effect on dissociation (Table VI). Of these only ADP had an effect, and this was small. In these experiments essentially all of the tRNA which dissociated from the complex was recovered as free tRNA. TABLE THE

VI

EFFECT

OF

DIFFERENT

PHOSPHATE

COMPOUNDS

RELATIVE

TO

ATP

ON

THE

tRiiA--E~ZYxE

COMPLEX

42 ,ug of enzyme and r.g yg of SaP-labeled tRNA were incubated in a final volume of 0.2 ml under conditions described in the legend to Fig. IA. After 5 min at o’, the concentration of phosphate compound indicated was added and the mixture was incubated an additional IO min at 0”. Centrifugaticn and assay were as described in Materials and methods. The per cent, tRNA, bound and free, was determined by planimetry. Addition

CO?ZC?%. (mM)

None ATP ADP AMP TTP Inorganic potassium triphosphate Adenosine tetraphosphate

2.5 2.5 2.5 a.7 2.5 2.5

RNA (94) in RNA--enzyme comP?er 84 ‘5 56 75 7S 82

85 Biochim.

Biophys.

Acta,

174 (1969) 458-475

i;

7

z 01 ‘0 e VI

3 Y

2 P

CONCENTRATIONS

OF

NUCLEOSIDE

TRIPHOSPHATE

ON

THE

COMPLEX

FORMED

BETWEEN

DNA AND

ENZYME

??

DNA

82 59 13 42 38

(%)”

:: 58

18 41

(%)

44 6

0.0 0.0

(%)

DNA

97 53 94

98

(O/o)

Free

Bound

Free

Bound

DNA’

6.25 mM Nucleotide

5 mM Nucleotide DNA

DNA

Bound

43 0.0

0.C

74 0.0

(%)

Nucleotide

8 mM

57 38

_. exclusive

0.0

26 53

(%)

~______ Free DNA

_

* Bound DNA represents the i*C-labeled T4 DNA present in the sucrose gradient fractions resent DNA-enzyme aggregates which sedimented to the bottom of the tube. * The per cent DNA is relative to the total counts added to the reaction mixture.

UTP CTP

None ATP GTP

Nucleotide addition

of the free DNA.

37 4

0.0

81 0.0

(%I

DNA

-

It does not rep-

63 44

19 47 0.0

(%)

IO mM Nucleotide Free Bowad DWA

T4 DNA under the conditions described in the legend to Fig. r-4, in a final volume RNA polymerase was added to sonicated Y-labeled of 0.2 ml. After 5 min at o’, a final concentration of 5, 6.25, 8, or IO mM nucleoside triphosphstc was added as indicated. In the experiment containing 5 mM nucleoside triphosphatc, 59 pg of enzyme were incubated with 3.65 ,ug of sonicated T4 DNA: in the cxpcriment containing 6.25 mM XT!?, 63 pg of the enzyme were incubated with 7.30 pg of DNA; in the experiment containing 8 mM ATP, 61 pg of cnzynre were incubated with 3.65 /~g of DNA; and, in the experiment containing IO mM ATP, 57 pg of enzyme were incubated with 3.65 pg of DNA. The mixture was incubated an additional ro min at o0 and then layered on 4.4 ml of a linear s-20 “/” sucrose density gradient plus the nucleoside triphosphatc which was containing: 50 mM Tris-HCI (pH 7.5), 2 mM MnCI,, 8 mM MgCl,, 0.033 mM of (NH,),SO( present in the incubation and was added in the same concentration. Centrifugation and assay were as described in Matevials and methods The amounts of sonicated T4 DNA, bound and free, were determined by planimetry.

b

VII

TABLE

THE EFFECT OF DIFFERENT

3 f

STUDIES WITH THE

Effect

RNA POLYMERASE 1

of wcleoside

triphosphates

471 on the sonicated

T4 DNA-enzyme

complex.

Higher concentrations of nucleoside triphosphates were required to dissociate the sonicated T4 DNA-enzyme complex than to dissociate the RNA-enzyme complex. Effects of the nucleoticles on the DNA-enzyme complex were z-fold: (I) to dissociate the DNA which then sedimented less rapidly as free DNA and, (2) to cause aggregation of the complexes with a more rapid sedimentation. Table VII illustrates the effect on the DNA-enzyme complex of preincubation with each of the four nucleoside triphosphates at concentrations of 5 to IO mM. As with the RNA-nzyme complex, it is clear that purine nucleosicle triphosphates caused the largest decrease in the amount of the complex. Unlike the case of the RNA-enzyme complex however, GTP appeared to have a greater effect than ATP, and CTP caused more dissociation than UTP. The reason why the percentages of 14C-labeled DNA recovered do not in some instances add up to IOO y/o is that the nucleoticles caused aggregation of the DNA, particularly when the concentration was high. This effect was greatest with GTP and least with LJTP. For the concentration ranges examined, dissociation of the DNXenzyme complex also required the addition of divalent cation. The effect of nucleoside triphosphates on the activity of the RNA polymerase. Active enzyme could not be recovered in the sucrose density gradient fractions of experiments in which a nucleoside triphosphate dissociated the DNA- or RNA-enzyme complex. Therefore, experiments were designed to examine the inactivation effects of the different nucleosicle triphosphates. The enzyme was preincubated with one nucleoside triphosphate at a IO mM concentration at oO, and the mixture was then diluted to give a final nucleotide concentration of 0.4 mM. The remaining three nucleosicle triphosphates plus calf thymus DNA were then added, and RNA synthesis was measured. The data in Table VIII show the percent of inactivation relative to a control in which DNA and enzyme were preincubated without nucleoticle. With a

TABLE

VIII

INACTIVATION PHATES

AT

OF A

E.

Cdi

CONCENTRATION

RNA

POLYMERASE OF

IO

DUE

To

PREINCUBATION

WITH

NUCLEOSIDE

TRIPHOS-

mM

Prewwubation. Mixtures, 0.1 ml in volume, containing 50 mM potassium maleate (pH 7.5), 2 mM MnCl,, 8 mM M&l,, 5.4 mM mercaptoethanol, 142 ,ug RNA polymerase, and IO mM of the nucleoside triphosphate indicated were preincubated at o” for I or 30 min. Incubation. A o.on-ml aliquot, containing 0.2 ,umole of the designated nucleoside triphosphate, was then added to a reaction mixture which contained: buffer divalent cations, and mercaptoethanol as in the preincubation, 0.2 mM [32P]UTP in the experiment in which CTP was present in the preincubation, 0.2 mM 3H-labeled CTP in all other experiments, and 0.4 mM of the remaining nucleoside triphosphates that were not present in the preincubation. RN.4 synthesk was started by the addition of 18 ,ug of calf thymus DNA either immediately following addition of the premcubation aliquot or after a further preincubation of 30 min at 0’. Incubations were for 5 min at 38”. The RN:\ in o..j-ml aliquots was precipitated, washed, and counted as described2. A’ucleotide triphosphaic

ATI’ GTI’ IJTP CTI’

9.6 99 1.6 4.9

98 99 ‘7 38

97 99 18 34 B&him.

Biophys.

Actn,

174 (1969) 458-47.5

I). D.

472 IO

mM concentration

of nucleoside triphosphate

ANTHONY

L’f a/.

present for I min, GTP was the most

effective inactivator, followed by ATP and then CTP (ref. 14). This order of effect was similar to that found for dissociation and aggregation of the DNA-enzyme complex (Table VII).

A 3o-min

preincubation

of the enzyme

in IO mM nucleoside

tri-

phosphate (Table VIII) resulted in more pronounced inactivation. When this 3o-min preincubation was followed by a further 3o-min incubation with the same nucleotide at 0.4 mM, and then addition of the other three nucleotides similar

results

were obtained

suggesting

that

to initiate

the inactivation

RNA synthesis,

cannot

be reversed

readily by lowering the nucleotide concentration. From these experiments, clear that high concentrations of nucleotides inactivated the RNA polymerase, that these concentrations also dissociated the enzyme-nucleic nucleotides were more effective than pyrimidine nucleotides nomena.

it is and

acid complex. Purine for both of these phe-

DISCUSSION

The RNA polymerase Two different

binding

is an enzyme with a number

sites for nucleotides

of different

binding

must be present for the formation

sites. of the

first phosphodiester bond. Preliminary evidence for different nucleotide Km values has been presented2. For the binding of the nucleic acid polymer to the enzyme, a third site is required. Evidence has been presented in this paper to support the hypothesis that either DNA or RNA can be bound at this site. The results of five different approaches to the characteristics of polymer binding are very similar when either DNA or RNA is used as the polymer.

First,

it has been noted that by increasing

the ionic strength,

the enzyme-polymer complex can be dissociated12,14,31, and that the behavior RNA is like that of DNA. Second, the binding of both RNA and DNA is better the absence

than in the presence

of divalent

metals.

Third,

a single

of in

ribonucleoside

triphosphate, in the presence but not in the absence of divalent cation, under certain conditions, is able to displace either DNA or RNA from the enzyme. Although the ordering of the effectiveness of nucleotides is not exactly the same with RNA as compared to DNA, in both cases the purines are more effective than the pyrimidines. Fourth,

data have been presented

to show that single-stranded

or sonicated

double-

stranded DNA can displace tRNA bound to the enzyme while tRNA can displace high molecular weight double-stranded DNA bound to the enzyme. Fifth, competitive kinetics are obtained when tRNA is used to inhibit the RNA polymerase reaction in the presence of varying DNA concentrations. Although none of these observations provides absolute proof that the same site binds either DNA or RNA, this interpretation of the data is most reasonable. It is clear from the data presented and that of others12,2g that a polymerase molecule of approx. 13 S can bind to a tRNA molecule. The stoichiometry reported here indicates that at least one RNA molecule can be bound per 13-S enzyme molecule. An order of displacement of nucleic acid polymers from the enzyme-polymer complex has been established. The order is: sonic-treated single-stranded DNA > (displaces) sonic-treated double-stranded DNA > high molecular weight singlestranded DNA > tRNA > high molecular weight double-stranded DNA. It should Biochim.

Biophys.

Acta,

174 (1969)

458-475

STUDIES WITH THE

RNA POLYMERASE. 1

473

be stressed that this is a qualitative phenomenon where approximately equal weights of two of the polymers were added to the enzyme, the mixture allowed to stand at o0 for a period of time, and then analyzed. The order of addition of the polymers under these conditions did not affect the outcome. Different types of binding sites may exist segments, and in the different polymers; 3’-hydroxyl end groups30, single-stranded specific areas in double-stranded DNA may all be binding sites. Since the concentration of such sites is unknown in a unit weight of any of the polymers used, the quantitative aspects of this displacement phenomenon would be difficult to evaluate. It should be noted that displacement implies some form of affinity of the enzyme when bound to one polymer for the second polymer. Multiple polymer binding sites would be a possible explanation for this which could be tested. The significance of the nucleoside triphosphate effects is not clear. At higher concentrations, all single nucleoside triphosphates cause dissociation of the RNAenzyme complex, with an order of effectiveness of purines greater than pyrimidines. DNA-enzyme complexes are also affected more by the purine than by the pyrimidine nucleotides. The DNA complexes are dissociated by each of the four nucleoside triphosphates. In even higher concentrations, GTP, ATP, and CTP cause aggregation of the DNA-enzyme complexes so that the sedimentation is more rapid than that of the DNA-enzyme complex without added nucleotide. In the concentrations tested, GTP caused the greatest degree of aggregation of the complex. This suggests that GTP either allows an enzyme molecule to bind to more than one DNA molecule or that it alters a bound enzyme molecule so that it can aggregate with other bound enzyme molecules*. It is likely that the ability of nucleoside triphosphates to alter the enzymepolymer complex is related to their ability to inactivate the enzyme. This relationfirst, that active enzyme could not be ship is supported by the observations, recovered after the dissociation of the enzyme-polymer complex by a nucleotide, second, that single nucleotides were able to inactivate the enzyme when tested directly, and third, that the purine nucleotides were more effective than pyrimidine nucleotides both in altering the enzyme-polymer complex and in inactivating the enzyme. The effect of nucleotides on the polymer-enzyme complex and on the enzyme activity may be due to an alteration in the protein conformation which under these conditions is irreversible. At lower nucleotide levels it is possible that a conformational change occurs which is reversible. The nucleotide effect on polymer binding and on enzyme inactivation is probably due to nucleotide binding at one of the ribonucleoside triphosphate sites involved in the synthesis of RNA rather than at some nonspecific site. In support of this is the fact that a number of compounds related structurally to the ribonucleoside triphosphates had no effect on the dissociation or inactivation. For the nucleotide dissociation effect, a divalent cation was required. Some of the data presented in this communication and obtained by others can be correlated. Purine nucleotides are utilized preferentially for initiation of RNA synthesis. This has been observed by analysis of the 5’-terminal nucleotide, which is predominantly GTP when single-stranded DNA is the template”, and also by kinetic ’ Dr. C. W. Wu has observed in preliminary experiments with the analytical ultracentrifuge that elevated levels of GTP caused an interaction of the 13-S molecules of the enzyme to form an aggregate which sedimented very rapidly so that no Schlieren pattern was observed. Riochzm.

Biophys.

Ada,

174 (1969)

458-475

Il. D. ANTHOXV

474

Ctal.

analys9. Purine nucleotide interaction with the enzyme can be observed by other means. The dissociation of the polymer-enzyme complex occurs more readily in the presence of purine than of pyrimidine nucleotides, and furthermore the inactivation of the enzyme in the absence of DNA occurs primarily with purine nucleotides and especially with GTP (ref. 14). These two phenomena which occur with relatively high concentrations

of nucleotides

observed

GTP is the most effective

that

are described

in this paper. STEAD AND JONESES have nucleotide

in the prevention

of the heat-

induced loss of the capacity of the polymerase to bind to DNA. Although GTP helped to preserve the binding capacity, the enzymatic activity was lost. The correlation then is clear between the effect of purine nucleotides single-stranded

DNA)

on initiation

and the effect of purine nucleotides

(GTP > ATP with

on enzyme

inactivation

(GTP > ATP) or enzyme-DNA binding capacity. This correlation suggests that there is a nucleoside triphosphate binding site on the RNA polymerase with which purine nucleotides

interact

preferentially

More direct studies of nucleotide

to pyrimidine

interaction

nucleotides.

with the enzyme are being pursued

to try to obtain direct evidence for preferential binding of the purine nucleotides. If this preferential binding is observed it would be of interest to know whether the site of binding is that occupied by the 5’-terminal nucleoside this site imparts any specificity to the initiation process.

triphosphate

and whether

ACKNOWLEDGMENTS This GM06075

investigation

was supported

and in part by National

wish to thank

lysodeikticus

Institutes

by National

Institutes

of Health

of Health Grant ~TI-GM~~.

Grant

The authors

Dr. G. BECKING for the calf thymus DNA, Dr. C. W. WV for the M. DNA and Dr. E. MELGAR for the T4 DNA.

REFERENCES I H. BREMER AND M. W. KONRAD, Proc. Natl.Acad. Sci.U.S.,51 (1964)801. 2 D. D. ANTHONY, E. ZESZOTEK AND D. A. GOLDTHWAIT, Proc. Natl. Acad. Sci. U.S., 56 (1966) 1026. 3 J. S. KRAKOW AND S. OCHOA, Proc. Natl. Acad. Sci. U.S., 49 (1963) 88. 4 C. F. Fox, W. S.ROBINSON, R. HASELKORN AND S. B. WEISS, J. Biol. Chem., z3g (1964) 186. 5 F. C. CLARK AND M. J. TAYSIR, J. Biol. Cham., 240 (1965) 3379. 6 S. K. NIYOGI AND A. STEVENS, J. Biol. Cham., 240 (1965) 2587. 7 A. TISSI&RES, S. BOURGEOIS AND F. GROS, J. Mol. Biol., 7 (1963) IOO. 8 J. S. KRAKOW, Biochim. Biophys. Acta, 72 (1963) 566. g H. BREMER, C. YEGIAN AND M. KONRAD, J. Mol. Biol., 16 (1965) 94. 10 C. F. Fox, R. I. GUMPORT AND S. B. WEISS, J. Biol. Chem., 240 (1965) 2101. II A. STEVENS, A. J. EMERY, JR. AND N. STERNBERGER, R&hem. Biophys. Res. Commun., 24 12 13 14 15 16 17 18 Ig 20 21

(1966) 929. J, P. RICHARDSON, J. Mol. Biol., 21 (1966) 83. J. S. KRAKOW, J. Biol. Chem., 241 (1966) 1830. D. D. ANTHONY, E. ZESZOTEK AND D. E. GOLDTHWAIT, Federation Proc., G. S. STENT AND C. R. FUERST, J. Gen. Physiol., 38 (1954) 441. D. FRASER AND E. A. JERREL, J. Biol. Chem., 205 (1953) 291. G. ZUBAY, J. Mol. Biol., 4 (1962) 347. J. D. MANDELL AND A. D. HERSHEY, Anal. Biochem., I (1960) 66. N. SUEOKA AND T. CHENG, J. Mol. Biol., 4 (1962) 161. R. 0. HURST, Can. J. B&hem. Physiol., 36 (1958) 1115. R. B. INMAN AND R. L. BALDWIN, .I. Mol. Biol.. 5 (1962) 172.

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Actn,

174

(1969)

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25 (1966)

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STUDIES WITH THE

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POLYMERASE.

475

1

22 IT. MAITRA AND J. HURWITZ, J. Biol. Chem., 242 (1967) 4897. 23 D. A. GOLDTHWAIT AND J. L. STARR, J. Biol. Chem., 235 (1960) 2025. 24 E. CHARGAFF AND J. N. DAVIDSON, The Nucleic Acids, Vol. I, Academic

Press, New York,

1955, P. 302.

25 K. BURTON, Biochem. J.. 62 (1956) 315. 26 1. P. RICHARDSON, Proc. Natl. Acad. Sci. U.S., 55 (1966) 1616. 27 b. A. SMITH, A.M.MARTINEZ, R.L. RATLIFF,D.-c. ~~LL;AMS AND R.N.HAYEs,

Biochemistry, 6 (1967) 3057. 28 0. W. JONES AND P. BERG, J. Mol. Biol., 22 (1966) Igg. 2g H. BREMER _&ND G. S. STENT, Acta Biochim. Polon., 13 (1966) 367. 30 P. BERG, R. D. KORNBERG, H. FANSCHER AND M. DIECKMANN, Biochem. Biophys. Res. Commun., 18 (1965) 932. 31 1'.CHAMBON, M. RAMUZ AND J. DOLY, Biochem. Biophys. Res. Commun., 21 (1965) 156. 32 E. J. FREEMAN AND 0. W. JONES, Biochem. Biophys. Res. Commun., 29 (1967) 45. 33 N. STERNBERGER AND A. STEVENS, Biochem. Biophys. Res. Commun., 24 (1966) 937. 34 I‘. MAITRA AND J. HURWITZ, Proc. Natl. Acad. Sci. U.S., 54 (1965) 815. 35 N. W. STEAD AND 0. W. JONES, Biochim. Biophys. Ada, 145 (1967) 679. Riochim.

Biophyr.

Acta,

174

(1969)

458-475